Published online March 12, 2007
Soil Microbial Fingerprints, Carbon, and Nitrogen in a Mojave Desert Creosote-Bush Ecosystem Stephanie A. Ewing Ecosystem Science Division ESPM Univ. of California Berkeley, CA 94720
Randal J. Southard* Soil Science Graduate Group Land, Air and Water Resources One Shields Ave. Univ. of California Davis, CA 95616
Jennifer L. Macalady Geosciences Dep. The Pennsylvania State Univ. University Park, PA 16802
Anthony S. Hartshorn
Abbreviations: DOC, dissolved organic carbon; PLFA, polar lipid fatty acid.
FOREST, RANGE & WILDLAND SOILS
Geography Dep. Univ. of California Santa Barbara, CA 93106
Creosote-bush [Larrea tridentata (Sessé & Moc. ex DC.) Coville] shrubs in California’s Mojave Desert support well-developed soil resource islands, where individual shrubs define areas of elevated soil nutrients, water-holding capacity, and microbial activity. To better understand the spatial variability of microbial communities and potential impacts on nutrient cycling in shrub ecosystems, we examined microbial communities using polar lipid fatty acids (PLFA) and several soil properties including δ15N, DNA, C and N contents under mature shrubs and as a function of horizontal distance (0–3 m) away from the base of the shrubs. Shrub-base soils (0 m) contained more C and N, were slightly more acidic, and supported significantly larger microbial populations than soils between shrubs. The PLFA fingerprints also suggested that microbial communities, particularly at the shrub base, had a different composition than soils between shrubs, including a higher proportion of actinomycetes containing the biomarker 10me17:0. Soil respiration was generally highest at 0 m, corresponding with larger microbial biomass and larger C and N pools, but was highly variable, probably due to contributions from grasses and forbs. Average δ15N values resembled plant material at the shrub base (4‰) and were significantly isotopically enriched away from the shrubs (7‰), suggesting that fractionating losses of soil N occurred between shrubs. The elevated nutrient status of resource islands supported soil microbial communities that were larger, were different in character, respired more actively, and cycled N more tightly than those found in open spaces between shrubs. These open spaces “leak” isotopically light N from the soil.
Mara J. Johnson Soil Science Graduate Group, Land, Air and Water Resources, One Shields Ave., Univ. of California, Davis, CA 95616
T
he patchy distribution of soil nutrients in shrub–desert ecosystems has been well characterized (Schlesinger et al., 1990), but less is known about how soil microbial community composition, microbial respiration, and soil N-cycling processes contribute to the “islands of fertility” that form under shrub canopies. These islands are locations where plant litter accumulates, where wind-borne material may be trapped under shrubs (Garcia-Moya and McKell, 1970; Schlesinger et al., 1996; Holzapfel and Mahall, 1999), and where the soil may be protected from raindrop impact, erosion, and runoff. Kieft et al. (1998) described distinct islands of C, N, and microbial biomass under L. tridentata in New Mexico and showed that temporal variation of these soil resources was greatest under L. tridentata shrubs, compared with soils under grasses in a nearby grassland biome or to soils in bare spaces in either the shrubland or grassland. Spatial heterogeneity of microclimatic conditions under shrubs vs. open spaces between shrubs has been documented in the cool, semiarid shrublands of eastern Utah (Forseth et al., 2001),
Soil Sci. Soc. Am. J. 71:469–475 doi:10.2136/sssaj2005.0283 Received 26 Aug. 2005. *Corresponding author (
[email protected]). © Soil Science Society of America 677 S. Segoe Rd. Madison WI 53711 USA SSSAJ: Volume 71: Number 2 • March–April 2007
where shrubs lowered soil temperatures and reduced soil water content, particularly in the upper 20 cm of the soil, compared with open microhabitats, although soil N, P, and organic matter showed little shrub island effect. Shrub-associated islands of fertility in the Mojave Desert of California, where some individual L. tridentata have existed for several thousand years (Vasek, 1980), are well developed in terms of soil nutrient variation (Schlesinger et al., 1996). This distinct patchiness makes the Mojave Desert a suitable location to explore the variation in microbial communities, soil respiration, and soil N isotopic composition that may accompany changes in N-cycling processes as a function of proximity to shrubs. Stable soil N isotopic signatures (δ15N) are a convenient tracer of net soil N-cycling effects because they represent the balance of biological N transformations (Amundson and Baisden, 2000). Previous work using soil δ15N has suggested less conservative cycling of N in hot, dry locations, where losses due to isotopefractionating processes result in higher soil δ15N signatures than at cooler, more moist sites (Amundson and Baisden, 2000; Austin and Vitousek, 1998; Riley and Vitousek, 1995; Schulze et al., 1991). In arid zones, processes that favor loss of 14N (and thus increase soil δ15N) may occur during brief episodes of denitrification following rainfall events; by NH3 volatilization, which is favored in dry, alkaline soils with low cation exchange capacity; or as loss of NOx and N2O during nitrification and denitrification (Mummey et al., 469
1997; Schlesinger et al., 1990; Smart et al., 1999). To the extent that N-cycling processes are a function of plant cover and climate, we hypothesized that these processes vary at the scale of microclimates beneath shrubs in desert ecosystems. Elevated soil microbial biomass is commonly observed under shrub canopies (Bolton et al., 1993; Smith et al., 1994; Sarig et al., 1999), but variations in the character of the microbial community with shrub proximity have not been sufficiently explored. Microbial community profiles are likely to vary with shifts in microclimate and N-cycling processes between resource islands and open spaces (Steinberger et al., 1999; Kinsbursky et al., 1990; Herman, 1997). The aim of this study was (i) to determine whether variation in the soil microbial community coincides with variations in soil respiration and net N cycling between resource islands and shrub interspaces, and (ii) to provide a preliminary characterization of the nature of any observed variation in the microbial community.
MATERIALS AND METHODS Field Site Fieldwork was conducted in March 1998 on the toe of an active alluvial fan north of Ludlow, CA (?250 km northeast of Los Angeles, 34.9° N, 116.2° W), near the Broadwell Lake playa at an elevation of 400 m. Local mean annual air temperature is estimated to be 22°C (hyperthermic soil temperature regime), and mean annual precipitation is estimated to be 100 mm (aridic soil moisture regime) based on elevation and plant community composition (Miles and Goudie, 1997). The slope at the study site is 2%, with a southeast aspect. Mesozoic granitic and Miocene volcanic rocks (Rogers, 1975) are the dominant sources for the sandy alluvium from which soils at the study site formed. Although creosote-bush is the dominant plant species, the local plant community also includes burrobrush [Ambrosia dumosa (Gray) Payne], brittlebush (Encelia farinosa Gray), and annual grasses and forbs.
Field Sampling We selected eight creosote-bush shrubs, approximately equally spaced around the perimeter of a circular area about 75 m in diameter. These shrubs were numbered 1 through 8 and were the primary focus of our sampling. We also obtained samples from eight additional shrubs around this perimeter for a few of the analyses described below. Three-meter-long transects were established north and east of each shrub. The maximum shrub diameter and the radius of each shrub along the north transect axis were measured. Shrub maximum diameters ranged from 1.7 to 4.0 m (average of 3.0 m). On each transect, we located sampling sites immediately adjacent to the main stem (0 m, which we refer to as “base” samples) and at horizontal distances of 1, 2, and 3 m from the base. For 14 of the 16 shrubs, the sampling sites at 1 m from the base were under the shrub canopy. Field measurements tested the effect of soil water content on soil respiration. We pushed 5-cmhigh by 10-cm-diameter polyvinyl chloride (PVC) rings about 1 cm into the surface soil at the transect sampling sites. On the north transects of four of the shrubs (Shrubs 2, 4, 6, and 8), we added 100 mL of water to each ring to simulate a rainfall event of about 1.3 cm, which we estimated would wet the soils to a depth of about 15 cm. The north transects of Shrubs 1, 3, 5, and 7 were not watered. After about 24 h, soil respiration was measured in the watered and unwatered rings using an EGM-1 infrared gas analyzer (PP Systems, Haverhill, MA), with the following operating conditions: chamber diameter, 100 mm; chamber volume, 1170 cm3; headspace sampled every 8 s, for 120 s final reading; fan speed, 300 to 350 cm3 min−1. We attempted to measure hydraulic conductivity of the soil at several locations with a constant-head permeameter, but hydraulic conduc470
tivity was too rapid to obtain reliable results. Based on this field experience and soil texture, we estimated the hydraulic conductivity to be in the 10 to 100 μm s−1 range (Soil Survey Division Staff, 1993). A soil pit was excavated near the base of one of the shrubs. The pedon was described and sampled by horizon for particle size analysis. In addition, soil samples were collected from the (approximately) upper 2 cm of soil within each PVC ring at each of the unwatered transect sampling sites (0, 1, 2, and 3 m), and immediately outside each PVC ring at the watered transect sampling sites for lab analyses. We assumed that microbial communities would be most concentrated very close to the soil surface in this hot, dry climate, so we focused our attention on the upper 2 cm for the majority of our analyses. The upper few centimeters of soils are also those most likely to be affected by localized erosion and deposition (e.g., Kieft et al., 1998). These samples, and samples from six additional shrubs with north and east transects (14 shrubs total), were transported in plastic bags to the lab at ambient temperature for analysis of field-moist water content and pH, carbonate content, total C and N content, and 15N content of air-dried soil. For DNA, PLFA, and soluble C and N analyses, an additional set of samples was collected from the upper 2 cm of soils at the unwatered north transect sampling sites (0, 1, 2, and 3 m) of Shrubs 1, 3, 5, and 7. These samples were transported on ice, stored frozen (−20°C), and processed frozen (not air dried).
Lab Analyses Particle size distribution of air-dried soil material was measured on the pedon samples by the pipette method (Soil Survey Staff, 1996). Unwatered samples from all 14 shrubs were analyzed as follows. Gravimetric water content of the field-moist samples was calculated by mass difference after 105°C oven drying overnight. Total C and N and δ15N in air-dry soil were measured using an Integra-CN integrated combustion, purification, and measurement system (Europa Scientific, Crewe, UK). Carbonate was measured in samples from north and east transects of Shrubs 1, 3, 5, 7, 9, and 13 by reacting 1 g of soil with 12 mL of 12 M HCl in a closed jar (similar to the method of Soil Survey Staff [1996]). The resulting CO2 gas was quantified using a Horiba infrared gas analyzer (Southeastern Automation Group, Knoxville, TN). Carbonate concentrations were calculated by comparing the evolved CO2 to standard curves prepared from CO2 gas standards of known concentration, as well as CO2 generated by combining analytical-grade CaCO3 with acid. Soil pH of all samples from Shrubs 1, 3, 5, and 7 was measured in 1:1 soil/ water mixtures with a glass electrode (Soil Survey Staff, 1996). Soluble C and N were measured in the frozen samples from north transects of Shrubs 1, 3, 5, and 7 (unwatered). Three replicate soil extracts were prepared from each sample by standard KCl extraction (Keeney and Nelson, 1982), and frozen (−20°C) for subsequent analysis. The KCl extracts were diluted 1:4 with ultrapure (18 Mω) water before analysis for dissolved organic C (DOC) using a Phoenix 8000 UV-persulfate C analyzer (Tekmar-Dohrmann, Cincinnati, OH). Dissolved NH4+ and NO3− in the extracts were measured colorimetrically using a QuickChem 8000 automated ion analyzer (Lachat Instruments, Milwaukee, WI). Frozen samples from the north transects of Shrubs 1, 3, 5, and 7 (unwatered) were analyzed for PLFA. Triplicate 5-g subsamples were extracted with a one-phase solvent extractant using a modification of the method of Bligh and Dyer (1959). Polar lipids (including phospholipids) were separated from neutral and glycolipids using solid-phase extraction columns (0.50 g Si, Supelco, Bellefonte, PA). The polar lipid fraction was subjected to mild alkaline methanolysis as described previously (Bossio and Scow, 1998), and the resulting fatty acid methyl esters (FAMEs) were extracted with two aliquots of hexane. The hexane was evaporated under N2 at room temperature SSSAJ: Volume 71: Number 2 • March–April 2007
SSSAJ: Volume 71: Number 2• March–April 2007
†† Values followed by the same letter are not significantly different (P < 0.05).
# DNA measured at Shrubs 3 and 7.
¶ Soil prewetted; respiration on wetting measured at Shrubs 2, 4, 6, and 8.
‡ CO3–C (carbon derived from calcium carbonate equivalents) and Q (gravimetric water content under field conditions) measured at 14 shrubs.
§ pH, CO2–C (carbon in measured flux of CO2) dry (soil not prewetted), PLFA (polar lipid fatty acid), DOC (KCl-extractable dissolved organic carbon), NH4+–N and NO3−–N (KCl-extractable), and Nmin (sum of NH4+ and NO3−) measured at Shrubs 1, 3, 5, and 7.
228 180 198 90 87 118 94 6 35 38 25 115 99 CV, %
† Total C, N, and δ15N calculated from transects from 14 shrubs; samples with N contents 99%, Table 1), but δ15N values were lower in base soils (4.1 ± 0.31‰, Table 1) relative to interspace soils (δ15N = 6.8 ± 1.34‰, Table 1), suggesting variation in N sources or, more likely, in transformation and loss of N. Base δ15N values were comparable to average values for L. tridentata leaves observed by Shearer et al. (1983) in the Sonoran Desert. If CO3–C is excluded from the calculation of C/N, the ratio tends to decrease from 0 to 3 m (compare with C/N values in Table 1), which suggests that the interspace soil organic matter was more highly decomposed than base soil organic matter. Thus, the enriched interspace δ15N values could reflect losses of isotopically light N during decomposition of L. tridentata litter. Alternatively, the C/N ratio may reflect variation in the quality of litter and rates of plant litter input under shrubs compared with spaces between shrubs. Notably, the degree of δ15N variation that we observed between base and interspace soils was similar to the variation observed between canopy and intercanopy soils associated with Juniperus osteosperma (Torr.) Little (Utah juniper) in southern Utah (Evans and Ehleringer, 1993). SSSAJ: Volume 71: Number 2• March–April 2007
Fig. 1. (a) Redundancy analysis of polar lipid fatty acid (PLFA) abundance from transects away from four shrubs (a, b, c, d). Distance from the shrub main stem is shown as 0, 1, 2, or 3. Each point represents three replicate PLFA analyses. Ellipses identify samples collected at equal distances from shrubs. (b) Redundancy analysis for PLFA variables. Fatty acids that lie close to samples in plot (a) above if plot origins were superimposed would probably have a high relative abundance in those samples.
Soil δ15N values may broadly reflect temperature and precipitation, as hotter and drier climates yield increased soil δ15N compared with cooler, wetter climates (Amundson and Baisden, 2000; Austin and Vitousek, 1998; Schulze et al., 1991). The N isotope differences (Table 1) may indicate that this correlation holds for the presumably cooler and slightly moister microclimate created by the shrub canopy. At this
Fig. 2. Dependence of δ15N on N concentration at 3 m from shrubs, the only location where the relationship was significant (P = 0.01). 473
Mojave Desert site, N concentration and δ15N were inversely correlated only in the samples at 3 m (R2 = 0.23, Fig. 2), and were not correlated at other locations under or between the shrubs. We interpret these results to indicate loss of isotopically light N from interspace soils, thereby enriching δ15N, but this interpretation warrants further investigation, in particular with respect to soil processes that might underlie our observations. Although our data did not reveal a specific mechanism of fractionating N loss, we speculate that several mechanisms are possible in this environment. Fractionating N loss may occur via NO3 leaching following incomplete nitrification (if, for example, low water potentials were to limit transport of NH4 to NH3 oxidizers and some NH4 would not be nitrified), or gaseous N loss through denitrification, NH3 volatilization, or nitrification (Austin and Vitousek, 1998; Schlesinger et al., 1990; Smart et al., 1999). Denitrification (loss of N2 and secondary N oxides) would have been an unlikely process at this site, given the rapid soil permeability (Peterjohn and Schlesinger, 1991), but other mechanisms of fractionating loss are possible. Ammonia volatilization may have occurred, given the neutral to alkaline soil pH values (Schlesinger et al., 1990). Interspace NO3 leaching would have been probable (Amundson et al., 1989), and nitrification would have been rapid given the favorable warm temperatures, neutral to alkaline pH, and coarse soil texture. Even if nitrification were not limited, NOx emission during nitrification may have occurred and may have fractionated N isotopes (Smart et al., 1999; Riley and Vitousek, 1995; Evans and Ehleringer, 1993). It has been argued that N and water are limiting to net primary productivity in the Mojave (Chew and Chew, 1965; Sharifi et al., 1988; Lajtha and Whitford, 1989). These limitations would not preclude leaching of N from surface soils during short, intense rain events, or during wetter-than-average years, particularly between shrubs, where biological demand for N is presumably lower than under shrubs. The presence of deep (several meters) NO3 reservoirs beneath some desert soils (Walvoord et al., 2003) indicates that deep N transport can occur in aridic climates, and this process is possible at our sites, especially between shrubs. One interpretation of our results is that N was more tightly cycled under wetter, cooler conditions close to shrubs, where plant uptake and growing microbial populations restricted N loss. Shrub radius correlated positively with PLFA (R2 = 0.65), soil moisture (R2 = 0.96), total soil N (R2 = 0.65), and NH4+ (R2 = 0.64) in base samples, suggesting a positive feedback in which shrubs enhanced resource islands as they grew (or vice versa), and that the effect was most intensely focused closest to the main stem. We speculate that in resource islands, immobilization of N during relatively wet, productive periods in spring, and mineralization of N during drier times, contributed N for relatively rapid plant uptake as the system transitioned to drought conditions. Because we did not attempt to control for the relative abundance of annuals in intershrub sampling areas, we could not quantify the extent to which annuals might have contributed to greater variability in respiration and N isotopes with distance from shrubs. Although shrubs may control the location of resource islands (Halvorson et al., 1995), competition with grasses (Holzapfel and Mahall, 1999; Caldwell et al., 1985) 474
and proliferation of roots where nutrients occur (Jackson and Caldwell, 1989; Robinson et al., 1999) could have contributed to tighter N cycling in nutrient patches. Future research to characterize microbial community variation in shrub ecosystems should account for plant competition and root plasticity controlling nutrient availability. In summary, variation in soil microbial community fingerprints accompanied variation in the distribution of soil respiration and C and N with proximity to Larrea tridentata shrubs in the eastern Mojave. Both PLFA and DNA indicated that microbial biomass was greatest beneath the shrubs, a location also characterized by maximum soil respiration rates. The PLFA analyses showed (i) that both base and interspace soil microbial communities contain actinomycetes, organisms that may hold a competitive advantage in regions of low water potential, and (ii) that the composition of the microbial community at the base of the shrubs and in the space between shrubs was different. Fractionating losses of N may dominate soil N inventories between shrubs, and may result in leaching of isotopically depleted N to deep soil reservoirs. The degree to which resource islands limit soil N loss in shrub ecosystems may be a function of how efficiently microbial populations in those islands capture leachable N during rain events. ACKNOWLEDGMENTS We thank DiG McGahan, Rebecca Neumann, Atac Tuli, Peter Brostrom, and Gary Weissman for field and lab assistance, and the UC Davis Soil Science Graduate Group for financial support. REFERENCES Amundson, R., and W.T. Baisden. 2000. Stable isotope tracers and mathematical models in soil organic matter studies. p. 117–137. In O.E. Sala et al. (ed.) Methods in ecosystem science. Springer, New York. Amundson, R.G., O.A. Chadwick, J.M. Sowers, and H. Doner. 1989. Soil evolution along an altitudinal transect in the eastern Mojave Desert of Nevada, U.S.A. Geoderma 43:349–371. Austin, A.T., and P.M. Vitousek. 1998. Nutrient dynamics on a precipitation gradient in Hawai’i. Oecologia 113:519–529. Bligh, E.G., and W.J. Dyer. 1959. A rapid method of total lipid extraction and purification. Can. J. Biochem. Physiol. 17:297–302. Bolton, H., Jr., J.L. Smith, and S.O. Link. 1993. Soil microbial biomass and activity of a disturbed and undisturbed shrub-steppe ecosystem. Soil Biol. Biochem. 255:545–552. Bossio, D.A., and K.M. Scow. 1998. Impacts of carbon and flooding on soil microbial communities: Phospholipid fatty acid profiles and substrate utilization patterns. Microb. Ecol. 35:265–278. Brereton, R.G. 1990. Chemometrics: Applications of mathematics and statistics to laboratory systems. Ellis Horwood, New York. Caldwell, M.M., D.M. Eissenstat, J.H. Richards, and M.F. Allen. 1985. Competition for phosphorus: Differential uptake from the dual isotopelabeled soil interspaces between shrub and grass. Science 229:384–386. Chew, R.M., and A.E. Chew. 1965. The primary productivity of a desertshrub (Larrea tridentata) community. Ecol. Monogr. 35:355–375. Digby, P.G.N., and R.A. Kempton. 1987. Multivariate analysis of ecological communities. Chapman and Hall, New York. Evans, R.D., and J.R. Ehleringer. 1993. A break in the nitrogen cycle in aridlands? Evidence from δ15N of soils. Oecologia 94:314–317. Forseth, I.N., D.A. Wait, and B.B. Casper. 2001. Shading by shrubs in a desert system reduces the physiological and demographic performance of an associated herbaceous perennial. J. Ecol. 89:670–680. Garcia-Moya, E., and C.M. McKell. 1970. Contribution of shrubs to the nitrogen and phosphorus economy of a desert wash plant community. Ecology 51:81–88. Halvorson, J.J., J.L. Smith, H. Bolton, and R.E. Rossi. 1995. Evaluating shrub-associated spatial patterns of soil properties in a shrub–steppe SSSAJ: Volume 71: Number 2 • March–April 2007
ecosystem using multiple variable geostatistics. Soil Sci. Soc. Am. J. 59:1476–1487. Herman, R.P. 1997. Shrub invasion and bacterial community pattern in Swedish pasture soil. FEMS Microbiol. Ecol. 24:235–242. Holzapfel, C., and B.E. Mahall. 1999. Bidirectional facilitation and interference between shrubs and annuals in the Mojave Desert. Ecology 80:1747–1761. Jackson, D.A. 1993. Multivariate analysis of benthic invertebrate communities: The implication of choosing particular data standardizations, measures of association, and ordination methods. Hydrobiologia 268:9–26. Jackson, D.A. 1997. Compositional data in community ecology: The paradigm or peril of proportions? Ecology 78:929–940. Jackson, R.B., and M.M. Caldwell. 1989. The timing and degree of root proliferation in fertile-soil microsites for three cold desert perennials. Oecologia 81:149–153. Kampfer, P., and R.M. Kroppenstedt. 1996. Numerical analysis of fatty acid patterns of coryneform bacteria and related taxa. Can. J. Microbiol. 42:989–1005. Keeney, D.R., and D.W. Nelson. 1982. N—inorganic forms. p. 643–698. In A.L. Page et al (ed.) Methods of soil analysis. Part 2. Chemical and microbiological methods. 2nd ed. Agron. Monogr. 9. ASA and SSSA, Madison, WI. Kieft, T.L., C.S. White, S.R. Loftin, R. Aguilar, J.A. Craig, and D.A. Skaar. 1998. Temporal dynamics of soil carbon and nitrogen resources at a grassland–shrubland ecotone. Ecology 79:671–683. Kinsbursky, R.S., R. Degani, G. Barnes, and Y. Steinberger. 1990. Rootmicrobial population dynamics in a soil profile under the canopy of the desert shrub Zygophyllum dumosum. J. Arid Environ. 19:261–267. Kroppenstedt, R.M. 1985. Fatty acid and menaquinone analysis of actinomycetes and related organisms. p. 173–197. In M. Goodfellow and D.E. Minnikin (ed.) Chemical methods in bacterial systematics. Soc. Appl. Bacteriol. Tech. Ser. 20. Academic Press, New York. Lajtha, K., and W.G. Whitford. 1989. The effect of water and nitrogen amendments on photosynthesis, leaf demography, and resource-use efficiency in Larrea tridentata, a desert evergreen shrub. Oecologia 80:341–348. Lechevalier, M.P. 1989. Lipids in bacterial taxonomy. p. 455–561. In W.M. O’Leary (ed.) Practical handbook of microbiology. CRC Press, Boca Raton, FL. Miles, S.R., and C.B. Goudie. 1997. Ecological subregions of California. Section and subsection descriptions. R5-EM-TP-005. USDA For. Serv. Pac. Southw. Region, San Francisco, CA. Mummey, D.L., J.L. Smith, and H. Bolton. 1997. Small scale spatial and temporal variability of N2O flux from a shrub–steppe ecosystem. Soil Biol. Biochem. 29:1699–1706. Olsson, P.A., I. Thingstrup, I. Jakobsen, and E. Baath. 1999. Estimation of the biomass of arbuscular mycorrhizal fungi in a linseed field. Soil Biol. Biochem. 31:1879–1887. Peterjohn, W.T., and W.H. Schlesinger. 1991. Factors controlling denitrification in a Chihuahuan desert ecosystem. Soil Sci. Soc. Am. J. 55:1694–1701. Rao, C.R. 1964. The use and interpretation of principal component analysis in applied research. Sankhya Ser. A 26:329–358. Ratledge, C., and S.G. Wilkinson (ed.). 1988. Microbial lipids. Academic Press, San Diego. Riley, R.H., and P.M. Vitousek. 1995. Nutrient dynamics and nitrogen trace gas flux during ecosystem development in montane rain forest. Ecology 76:292–304. Robinson, D., A. Hodge, B.S. Griffiths, and A.H. Fitter. 1999. Plant root proliferation in nitrogen-rich patches confers competitive advantage.
SSSAJ: Volume 71: Number 2• March–April 2007
Proc. R. Soc. Lond. Ser. B 266:431–435. Rogers, T.H. 1975. Geologic map of California, San Bernardino sheet, 1:250 000. Calif. Div. of Mines and Geol., Sacramento. Sarig, S., A. Fliessbach, and Y. Steiberger. 1999. Soil microbial biomass under the canopy of coastal sand dune shrubs. Arid Soil Res. Rehabil. 13:75–80. Schlesinger, W.H., J.A. Raikes, A.E. Hartley, and A.F. Cross. 1996. On the spatial pattern of soil nutrients in desert ecosystems. Ecology 77:364– 374. Schlesinger, W.H., J.F. Reynolds, G.L. Cunningham, L.F. Huenneke, W.M. Jarrell, R.A. Virginia, and W.G. Whitford. 1990. Biological feedbacks in global desertification. Science 247:1043–1048. Schulze, E.-D., G. Geauer, H. Ziegler, and O.L. Lange. 1991. Estimates of nitrogen fixation by trees on an aridity gradient in Namibia. Oecologia 88:451–455. Sharifi, M.R., F.C. Meinzer, E.T. Nisen, P.W. Rundel, R.A. Virginia, W.M. Jarrell, D.J. Herman, and P.C. Clark. 1988. Effect of manipulation of water and nitrogen supplies on the quantitative phenology of Larrea tridentata (creosote bush) in the Sonoran Desert of California. Am. J. Bot. 75:1163–1174. Shearer, G., D.H. Kohl, R.A. Virginia, B.A. Bryan, J.L. Skeeters, E.T. Nilsen, M.R. Sharifi, and P.W. Rundel. 1983. N2–fixation from variation in the natural abundance of 15N in Sonoran Desert ecosystems. Oecologia 56:365–373. Smart, D.R., J.M. Stark, and V. Diego. 1999. Resource limitations to nitric oxide emissions from a sagebrush–steppe ecosystem. Biogeochemistry 47:63–86. Smith, J.L., J.J. Halvorson, and H. Bolton, Jr. 1994. Spatial relationships of soil microbial biomass and C and N mineralization in a semi-arid shrub– steppe ecosystem. Soil Biol. Biochem. 26:1151–1159. Soil Survey Division Staff. 1993. Soil survey manual. Agric. Handbk. 18. U.S. Gov. Print. Office, Washington, DC. Soil Survey Staff. 1996. Soil survey laboratory methods manual. Soil Surv. Invest. Rep. 42. Version 3.0. Natl. Soil Surv. Center, Lincoln, NE. Soil Survey Staff. 1999. Soil taxonomy—A basic system of soil classification for making and interpreting soil surveys. 2nd ed. Agric. Handbk. 436. U.S. Gov. Print. Office, Washington, DC. Steinberger, Y., L. Zelles, Q.Y. Bai, M. Von Luetzow, and J.C. Munich. 1999. Phospholipid fatty acid profiles as indicators for the microbial community structure in soils along a climatic transect in the Judean desert. Biol. Fertil. Soils 28:292–300. Vasek, F.C. 1980. Creosote bush: Long-lived clones in the Mojave Desert. Am. J. Bot. 67:246–255. Verdonschot, P.F.M., and C.J.F. Ter Braak. 1994. An experimental manipulation of oligochaete communities in mesocosms treated with chlorpyrifos or nutrient additions: Multivariate analyses with Monte Carlo permutation tests. Hydrobiologia 278:251–266. Walvoord, M.A., F.M. Phillips, D.A. Stonestrom, R.D. Evans, P.C. Hartsough, B.D. Newman, and R.G. Striegl. 2003. A reservoir of nitrate beneath desert soils. Science 302:1021–1024. White, D.C., H.C. Pinkart, and D.B. Ringelberg. 1996. Biomass measurements: Biochemical approaches. p. 91–101. In Manual of environmental microbiology. Am. Soc. Microbiol. Press, Washington, DC. Zar, J.H. 1999. Biostatistical analysis. 4th ed. Prentice Hall, Upper Saddle River, NJ. Zelles, L. 1999. Fatty acid patterns of phospholipids and lipopolysaccharides in the characterisation of microbial communities in soil: A review. Biol. Fertil. Soils 29:111–129. Zhou, J., M.A. Bruns, and J.M. Tiedje. 1996. DNA recovery from soils of diverse composition. Appl. Environ. Microbiol. 62:316–322.
475 475
